The fields of flow cytometry, ion-channel electrophysiology, single cell electroporation, controlled shear force in vivo-simulating cell culture and numerous related biotechnology approaches stand to benefit from advances in the design of microfluidic devices for manipulation of cells and attendant apparatus.
Flow cytometry is a widely used technique for the counting and classification of single cells (Cottingham 2005). Because of its high throughput (1000s of cells/s) and reliability, it has become the most widely used method for cell population identification. It has far reaching applications, from pharma and academic research to drug cell based screening and cell line QC. In the diagnostic market it is used primarily for the quantification of blood cell content, and used to monitor RBC counts as well as counts for all of the major WBC types (blood panel).
While flow cytometry was originally based on electrical resistance changes in a flow capillary, almost all modern flow cytometers are now based on a laser for excitation and PMT for the detection of fluorescent signals from each cell, thus providing data which is multiplexed with straight cell count data. This allows for the identification and counting of a number of different cell types (through the use of fluorescent probes), as well as establishing relations between the fluorescence intensities recorded.
A key drawback of this technique is the loss of sub cellular information. Cell morphological parameters, as well as localization of fluorescent signals w/in the cell and the correlation between various stains are all lost in a process which simply integrates the fluorescence intensity over the whole cell and outputs one number per cell.
Until recently, this type of information has only been accessible through the use of Image Scanning Cytometry, basically a microscope on an XYZ stage that scans a thin cavity filled w/cellular suspension. This is an automation of the manual hemocytometer. This technique is significantly slower as compared to flow cytometry, and requires automatic focus and due to movement of the substrate in the XY plane. An additional drawback of this technique is the inability to sort cells.
Recently introduced imaging flow cytometers aim to combine the speed and ease of use of flow cytometry with the high information content of Image Scanning Cytometry (Bonetta 2005). The instrument images cells as they flow by at high velocity in a single file. The requirement of assembling in-flight images of single cells requires a great deal of custom technical development, which translates in a relatively high price for such devices (approx. $300 k for ImageStream 100) and relegates their use to core labs and pharmaceutical companies.
Ion channels are functional units of all living systems and fulfill a number of roles, from fast signal transmission in the nervous system to regulation of biochemical pathways. In turn, a number of disorders have been linked to ion channel malfunction. Ion channels are implicated in mental disorders such as Alzheimer's and epilepsy, as well as heart disease, diabetes and neuromuscular disease [Shaffer]. Consequently, these transmembrane proteins are attractive drug targets and constitute about 20-30% of new drug development campaigns [Southhan].
The electrophysiology recording technique, termed patch clamp, has emerged as the gold standard in the study of ion channel function. It is based on the ability to perform recordings of transmembrane currents through a specific ion channel type. Traditionally, patch clamp recording is accomplished with a micromanipulator-positioned glass pipette under a microscope [Sackmann]. This technique was perfected in 1981-'83 by Nether and Sackmann through the achievement of high resistance seals between the glass pipette tip and the cell membrane. The basic setup is illustrated in FIG. 5A. Current that passes through the ion channels in either the membrane patch or the whole cell membrane is recorded at different bias voltages.
In June 2005, the FDA mandated that all drugs must be tested against the potassium (K) ion channel hERG, whose unpredicted adverse modulation by several blockbuster drugs has been implicated in long-QT syndrome and subsequent sudden death by heart malfunction [Denyer]. It has been estimated that approximately 25-40% of all lead compounds show hERG activity in vitro [Bennett].
The gold standard assay for hERG safety screening is the patch clamp: the cell is voltage clamped in whole-cell configuration (using a glass pipette) while the test compounds are introduced extracellularly. The response of the cell to the test compounds is evident from the current response of the cell when the compounds have reached the ion channel's binding site which, in the case of hERG, is on the interior portion of the cell (see FIG. 6). FIG. 6 shows an outward rectifying potassium channel. As indicated in the diagram, the candidate hERG blocker (e.g., a compound to be tested) binds at the intracellular side of the molecule. Source: Enal Razvi [Southhan]. The problem with this approach is that the compounds are notoriously slow to diffuse through the membrane to reach this binding site (t>20 minutes). In addition, multiple compound concentrations need to be applied sequentially to the same patched cell in order to provide consistent measurement. Consequently, hERG measurements often fail due to the lack of long term stability for the high resistance seal between the glass pipette and the cell (seals often degrade in t<20 minutes).
Despite constant improvement of the traditional patch clamp technique, it remains laborious, requiring pipettes to be placed in the cell vicinity by a skillful operator using a micromanipulator under a microscope. Consequently, the patch clamp technique has been difficult to use in drug development, where high-throughput automated measurements are required. An automated patch clamp setup for high-throughput measurements using disposable devices would eliminate the prohibitive time investment of the traditional patch clamp, while maintaining its advantages over indirect measurements of ion channel behavior. The first approach to automated patch consisted of an array of robotically operated patch clamp pipettes (Axon, Inc.), to be used with large cells (Xenopus oocytes). The most serious drawbacks of this approach its inability to work with mammalian cell lines as well as the complexity of the manipulation system, while savings in terms of reagent use are minimal. A microfabricated patch clamp approach, if perfected, would solve both these problems. Currently available automated electrophysiology devices are employed by large organizations at large capital expense (greater than $400,000 per instrument) as well as a large cost per data point (about $10 per cell trap). They also retain important limitations in the area of optical observation and compound perfusion.
RNA interference is arguably the most powerful second-generation functional genomics technology currently available [Klemic]. Its high robustness, specificity, and efficacy in silencing targeted genes suggests its potential to father the development of a whole new class of drugs for an incredibly broad range of diseases. Before this can happen, however, significant challenges with respect to short interfering RNA (siRNA) delivery and targeting must be overcome.
One way to traverse the cell membrane and access the cell's interior is by temporarily increasing the permeability of the cell membrane. This can be accomplished via electroporation, a technique which uses high electric fields to induce structural rearrangements of the cell membrane. Pores result when the transmembrane potential exceeds the dielectric breakdown voltage of the membrane (0.2-1.5V) [Weaver]. Polar substances otherwise impermeant to the plasma membrane (such as dyes, drugs, DNA, proteins, peptides, and amino acids) can thus be introduced into the cell.
In the early 1980s, Eberhard Neumann et al. demonstrated the feasibility of electroporation for delivering DNA to a population of mammalian cells [Lundqvist]. Since then, this method of bulk electroporation has become a standard technique routinely used to simultaneously transfect millions of cells in culture. Most commercially available electroporation systems still use Neumann's approach without too much variation. Bulk electroporation requires very high voltages (kVolts) and has little control over the permeabilization of individual cells, resulting in suboptimal parameters. Moreover, because different cell types require different electric field parameters to electroporate, the system has to be calibrated to determine appropriate pulse conditions a priori without any real-time control. Reversible electroporation, in which the pores can reseal, is therefore difficult [Chang, D. C.]. As a result, most commercial systems focus on improving buffer solutions to improve cell viability. Examples of commercial electroporation platforms include the Gene Pulser Xcell™ Eukaryotic System (Bio-Rad Laboratories), BTX® HT 96 Well Electroporation System (BTX® Molecular Delivery Systems), Nucleofector™ 96-well Shuffle System (Amaxa Biosystems), and Axoporator 800A (Molecular Devices).
Single cell electroporation obviates many of the challenges associated with bulk electroporation but is less common. Lundqvist et al first demonstrated single cell electroporation using carbon fiber microelectrodes in 1998 [Lundqvist]. To induce electroporation, they placed microelectrodes 2-5 microns away from adherent progenitor cells. Other single cell electroporation techniques developed since include: electrolyte-filled capillaries [Nolkrantz (Electroporation)], micropipettes [Hass, Rae], and chips [Huang]. For successful single cell electroporation, the cell must either be isolated or the electric field well focused to target a particular cell [Nolkrantz (Functional Screening)]. Currently single cell electroporation is performed using laborious manual setups.
Contact adhesion between cells and surfaces, both inside and outside organisms, are central to a large number of biological phenomena. Some examples are blood clotting, tissue repair, immune and inflammatory response, bacterial infections, and cancer progression. A widely used method to quantify cell adhesion has been the application of a range of shear forces in flow chambers. The same methods are used to determine the cellular response to shear stress through mechanotransduction pathways.
The bulk of this type of research is currently performed using macroscopic laminar flow chambers. Current practice suffers from limited throughput, cumbersome apparatus assembly, experiment failure (i.e. by bubble introduction), and a limited range of applicable shear forces.
Thus, there remains a considerable need for alternative designs of microfluidic devices for manipulation of cells to support flow cytometry, cell ion-channel electrophysiology, single cell electroporation, controlled shear force in vivo-simulating cell culture and related technologies. The present invention satisfies these needs and provides related advantages as well.